Mapping fast DNA polymerase exchange during replication

Despite extensive studies on DNA replication, the exchange mechanisms of DNA polymerase during replication remain unclear. Existing models propose that this exchange is facilitated by protein partners like helicase. Here we present data, employing a combination of mechanical DNA manipulation and single fluorescent protein observation, that reveal DNA polymerase undergoing rapid and autonomous exchange during replication not coordinated by other proteins. The DNA polymerase shows fast unbinding and rebinding dynamics, displaying a preference for either exonuclease or polymerase activity, or pausing events, during each brief binding event. We also observed a ‘memory effect’ in DNA polymerase rebinding, i.e., the enzyme tends to preserve its prior activity upon reassociation. This effect, potentially linked to the ssDNA/dsDNA junction’s conformation, might play a role in regulating binding preference enabling high processivity amidst rapid protein exchange. Taken together, our findings support an autonomous replication model that includes rapid protein exchange, burst of activity, and a ‘memory effect’ while moving processively forward.

1.The authors should explain why exonuclease and polymerase activities are observed at high and low forces, respectively?2. What is the rationale behind 60% labeling of polymerase? 3. Are DNA polymerase trajectories shown in the kymographs continuous?The overlay of the junction trajectory makes it difficult to visualize.4. Are DNA polymerase always bound at the replication junction during the experiments? 5. Why do the authors use fairly high concentration of DNA polymerase, i.e., 30 nM for single-molecule fluorescence experiments?How does it impact the signal to noise ratio?Wouldn't the high polymerase concentration affect data interpretation while correlating the optical tweezers data and fluorescence signal?6.Why the signal to noise ratio is low in the kymographs?Similar studies suggest a high signal to noise ratio can be achieved (Li, S., Wasserman, M.R., Yurieva, O. et al., Nat.Commun.13, 2022; Lewis, Jacob S., et al., Mol.Cell 77, 2020).7. What does the red box in Figure 1E represent?8.In the kymographs (Figs.1E, 3A, 4D and others), what does the high intensity at the bottom and top represent?In Figs.4A and 4B, the authors indicate that the top and bottom intensity bands are attributed to signals from bead? Can authors clarify it and mention it in all the kymographs?9. Figure 1, line 36: … while the gray line represents fluorescence signal at the ssDNA/dsDNA junction.There is no gray line in Figure 1F? 10.What does the green trace at the back of the yellow trace in Fig. 1F represent?11.The authors should mention the total length of the DNA handle in the main text.12. Lines 202-204: This sentence should be used somewhere in the beginning to help authors understand the increasing/decreasing basepairs trends in Figure 1 and others.

Drs. Avinash Kumar and Yongli Zhang
Reviewer #2 (Remarks to the Author): This work combines simultaneous confocal fluorescence and force extension techniques to track the activity of bacteriophage T7 DNA polymerase.Both exonuclease and polymerase activity are explored, where the T7 polymerase is labeled with a fluorophore.As in standard force spectroscopy experiments, changes in the fraction of single and double stranded DNA are tracked through the change in the overall length of the template as one is converted into the other.The new technique revealed in this work directly correlates length changes to the location of the T7 as imaged confocally and in kymographs.This enables a careful exploration of the rates of T7 stalling, release and even replacement on the template.While T7 polymerase activity has been widely characterized in force extension experiments, simultaneous imaging and force facilities answering questions that cannot be answered by force spectroscopy alone, including how long each polymerase is bound and active.
Careful study of pauses and bursts of activity reveal that T7 binding and activity is surprisingly stochastic.The same polymerase may rapidly switch between active and inactive states, followed by release.The picture suggested by this data is not of a single, highly processive polymerase, but of a very rapid release and replacement by others from solution.Furthermore, many T7 appear to bind and never initiate polymerase/exonuclease activity.Together this indicates a 'memory' effect, where active polymerase/exonuclease activity is followed by more of the same and switching between the two modes is rarely seen.An intriguing explanation for this is proposed: that in solution, T7 exists in two conformations (and rarely switching), and unbinding T7 leaves the DNA junction in a specific state favorable only to T7 of the same conformation.This is very interesting work that represents a significant advance over previous polymerase experiments and may suggest similar properties for polymerases from other systems.Furthermore, the data appears to be of high quality (though only a single fluorophore is tracked, and labeling is only 60% efficient), and is carefully analyzed.Finally, several controls for photobleaching and diffusion along single and double stranded DNA are shown.With minor clarifications, this paper should be suitable for publication.

Questions:
Figure 1 and Methods: Were the tagged T7 filtered after attachment? Figure 1 and Methods: How large is the tag compared to the polymerase? Figure 1 and Methods: What is the temporal resolution of the experiment?The pixel width is ~1 sec, but the methods and results mention that only events of 3 pixels were considered.So only pol exchanges that take longer that 3-5 sec can be seen?This should be discussed in detail.
Figure 1: The red box is not explained in the caption -is it for background subtraction?Figure 2: There are a high percentage of non-fluorescent events that show exonuclease activity.Could any of this be due to DNA peeling at these high forces over these long times?For all figures, please check the panel ordering.It is confusing to follow panels that go in order right to left, then down, then back up to the top row.
The first equation is equation 3, but shouldn't this be started at 1? Reviewer #3 (Remarks to the Author): The authors present single molecule kinetic analysis of DNA polymerization by monitoring changes in length under tension on the template strand resulting from differences between single strand and double stranded DNA.By simultaneously monitoring fluorescence of labeled enzyme and polymerization, the authors describe exchange between polymerases during polymerization.The methods used by the authors are clever and the data appear to be interpreted rigorously.However, their method is unable to resolve single nucleotide addition steps and is limited to 10 ms per time point.It is not the "high resolution" the authors claim.Moreover, the force on the DNA alters the kinetics as described below.
There is no description of the formation of a replication fork.The T7 DNA polymerase replication complex has been well characterized defining the coupled action of the helicase, polymerase, and exonuclease with leading and lagging strand synthesis (1).Rather it appears that the authors are only examining DNA elongation with a template and a primer with no duplex DNA and helicase ahead of the polymerase.The current study ignores these detailed studies and presents data that conflict with wellestablished properties of the polymerase.
The most severe limitation of the paper is the lack of quality controls for the changes induced by modification of the DNA polymerase.Most importantly, the authors use T7 gp5 polymerase fused with thioredoxin accessory protein.The only reference given in the text to the use of the fusion protein is a 1987 paper from Richardson's lab, which did not use a fusion protein.Studies have shown that without thioredoxin the DNA polymerase is no longer processive and "undergoes frequent dissociation from and rebinding to the DNA" (2), similar to the result reported in the current manuscript.No controls are given in this manuscript to assess the effect of fusing the thioredoxin to the polymerase.This is a fatal flaw since thioredoxin binding greatly affects polymerase activity and DNA dissociation rates.Fusing thioredoxin to achieve optimal interactions with the flexible DNA binding domain of the polymerase is not a trivial task and would require significant refinement and testing by established methods to define polymerization rates and processivity.A likely explanation of the results presented in this paper is that a less than optimal covalent linkage of thioredoxin to the polymerase was insufficient to stabilize an otherwise flexible DNA-binding domain leading thereby failing to achieve the desired tight DNA binding.
The expression of the T7 pol-thioredoxin fusion protein with an additional SNAP tag (182 amino acids, 19.4 kDa) is also not well described.Was the SNAP tag added to the N-or C-terminus of T7 gp5 and was a flexible linker included?The SNAP tag is likely to introduce a significant change to the polymerase structure and dynamics for which there are no controls.The addition of a large fluorophore further compounds problems.T7 DNA polymerase is a dynamic enzyme where conformational changes dictate nucleotide specificity and proofreading.T7 DNA polymerase has been extensively characterized by single turnover kinetic methods with single base-pair resolution, far exceeding the capabilities of single molecule methods to define reaction kinetics and mechanism (3-9).The results of numerous studies are at odds with the authors' conclusions.The enzyme catalyzes replication at ~300 bp/s and dissociates from DNA at with a rate constant of 0.2/s giving a processivity of 1500 bp.The exonuclease reaction on ssDNA occurs at 1000/s but with duplex DNA the exonuclease is governed by the rate of transfer of the primer strand from the polymerase to the exonuclease site.The kinetic partitioning between polymerization and exonuclease reaction has also been well defined.These studies should stand as a benchmark for evaluating any labeled enzyme to quantify the effects of modifications due to fusing with other proteins and adding fluorescent labels.
For each of these issues, the authors must address the effects of their significant enzyme modifications.Otherwise, their results only apply to the modified enzyme and are not applicable to polymerase function in the cell.At the very minimum, the following experimental measurements are necessary.Single turnover kinetic studies in solution can be easily performed with native and modified enzymes to quantify the effects of modifications.a.What is the active site concentration of the modified, labeled enzyme?That is, what fraction of the enzyme still binds and extends DNA? b.What is the rate of polymerization in solution?c.What is the Kd for DNA binding to the altered enzyme?d.Most importantly, what is the rate constant for DNA dissociation?
The effects of force on the DNA may be responsible for some of the unusual behavior seen in this report.Structural studies show that the template DNA enters the active site at 90 degrees to the growing duplex (10).Therefore, as suggested in the present studies and many previous reports using this method, adding tension to the two ends of the DNA distorts the polymerase active site.It is remarkable that the authors show that a force of 40-50 pN disrupts the polymerase activity so that only exonuclease reaction is seen.But they assume that dropping the force only 2-4-fold to 10-20 pN allows measurement of polymerization without affecting fundamental kinetic parameters including the polymerization and DNA dissociation rate.The authors' analysis rests on an unsupported assumption, and it is likely that the unique conclusions put forth by the authors regarding fast dissociation of the polymerase are an artefact of their methods and enzyme preparation, and there are no controls offered to overcome this objection.
Memory effects have been proposed from single molecule kinetic studies, but to my knowledge there has never been definitive data to support such postulates.Enzymes do not have memories; they are governed by rate constants for transitioning between states.

Response to Reviewers' Comments
Reviewer #1 (Remarks to the Author): In this manuscript, Xu et al. report the dynamics of T7 polymerase during DNA replication using correlative optical tweezers fluorescence microscopy.They track the DNA extension changes as the polymerase performs DNA polymerization and cleavage, as well as binding and unbinding to the DNA.They show that the polymerase acts in a stochastic and auxiliary proteins-independent manner at the ssDNA/dsDNA junction, where it carries out exonucleolysis and polymerization.They also provide experimental evidence of the memory effect that contributes to the polymerase processivity.The authors employ a novel assay to investigate the polymerase dynamics and reveal the correlation between the forcedependent polymerase activity and the fluorescence signal.The manuscript is well-written and presents findings that are of broad interest for researchers in the field.The manuscript meets the standards of the journal and deserves publication after some revisions.
1.The authors should explain why exonuclease and polymerase activities are observed at high and low forces, respectively?
We appreciate the reviewer's query regarding the force-dependent activities of DNA polymerase.The observed phenomena stem from the mechanical forces affecting the structure of the nucleic acids.At higher forces (i.e.: ~50 pN), the DNA molecule is stretched, which can expose the ends of the DNA strands, making them more accessible to exonucleases, removing nucleotides from the ends of DNA strands.This is crucial for removing damaged or mismatched nucleotides.Conversely, at lower forces (i.e.: ~ 20 pN), the DNA structure is more relaxed, allowing polymerases to effectively match and add nucleotides to the growing strand.Low mechanical stress ensures accurate base pairing and efficient strand elongation.This force-dependent behavior has been reported regularly over the last two decades, see for example these earlier studies on T7 DNA polymerase [1][2][3] .We now give brief background and suitable references about this in Methods section, with changes marked in red.

What is the rationale behind 60% labeling of polymerase?
The choice of a 60% labeling ratio for the polymerase was strategic to allow effective visualization of protein exchange at the replication fork.This incomplete labeling approach ensures that not every polymerase molecule is fluorescent, enabling the detection of new, unlabeled polymerases replacing labeled ones, thereby providing direct evidence of polymerase exchange dynamics.
3. Are DNA polymerase trajectories shown in the kymographs continuous?The overlay of the junction trajectory makes it difficult to visualize.
We acknowledge the reviewer's concern about the clarity of kymographs.The trajectories of DNA polymerase in the kymographs are not continuous due to the dynamic nature of polymerase binding and unbinding and the incomplete labelling.We now added the raw kymograph without the overlay as a panel in Figure S2A for clarity.4. Are DNA polymerase always bound at the replication junction during the experiments?During our experiments, DNA polymerase is not continuously bound at the replication junction.Our findings demonstrate that the polymerase undergoes rapid binding and unbinding events, contributing to the dynamic and stochastic nature of the replication process.This can also be observed by quantifying the fluorescence intensity along the ssDNA/dsDNA junction (Figure 1F and 1G).
In addition to binding at the ssDNA/dsDNA junction, DNA polymerases were also observed to bind on dsDNA, potentially searching for binding sites (Figure 4A).Finally, DNA polymerases were observed to bind stationary on ssDNA (Figure 4B).

5.
Why do the authors use fairly high concentration of DNA polymerase, i.e., 30 nM for single-molecule fluorescence experiments?How does it impact the signal to noise ratio?Wouldn't the high polymerase concentration affect data interpretation while correlating the optical tweezers data and fluorescence signal?
The 30 nM concentration of DNA polymerase was optimized and chosen to ensure a sufficient frequency of binding events and continuous activity for robust statistical analysis, following earlier studies 1,3 .In our experiments, a lower concentration of DNA polymerase could lead to reduced apparent processivity, potentially making it more challenging to detect these events.And importantly, we use confocal microscopy for imaging the DNA which provides a good signal to noise ratio because it images the fluorescent proteins that are on the DNA template while rejecting most of the background.
We appreciate the reviewers for their query regarding the signal-to-noise ratio (SNR) in our kymographs.It's important to note that the mentioned two publications investigate DNA polymerase systems that appear to have long lifetimes on the DNA; moreover, they were able to use different fluorophores with distinct imaging parameters.The SNR in our experiments is influenced by several factors, including the fluorescence intensity of labeled proteins (dye quantum yield), and the dynamic nature of polymerase interactions with DNA.
In our experiments, we optimized labeling to achieve a balance between detectable signal and maintaining the native functionality of the polymerase, as well as to observe the dynamics of DNA polymerase.In our earlier trials, over-labeling or using bright fluorophores (such as Atto647N) appeared to alter the protein dynamics.In addition, we think that the rapid and transient interactions of DNA polymerase with the DNA substrate can leads to fluctuating signal intensities.We now clarify this point more explicitly in the Method section.
7. What does the red box in Figure 1E represent?The red box in Figure 1E is utilized for background subtraction in our analysis.It defines a specific region in the kymograph where the background fluorescence signal is measured.This background signal is then subtracted from the fluorescence signal of the DNA polymerase to enhance the accuracy of our signal-to-noise ratio and ensure that the fluorescence changes we observe are indeed due to the binding and activity of the DNA polymerase, not background fluorescence.We clarified this in the figure caption.The high-intensity bands at the top and bottom of the kymographs are indeed due to fluorescence signals from the beads used in our optical tweezer's setup.These signals are not related to the DNA polymerase activity and serve as reference points in the kymographs.This information is now clearly mentioned in all relevant figure captions for better understanding in the revised manuscript.9. Figure 1, line 36: … while the gray line represents fluorescence signal at the ssDNA/dsDNA junction.There is no gray line in Figure 1F?
We apologize for the confusion.The mentioned gray line is the raw data of fluorescence signal of DNA polymerase at the ssDNA/dsDNA junction.To ensure the clarity, we revised the caption to "while the yellow line represents the filtered fluorescence signal of DNA polymerase at the ssDNA/dsDNA junction (raw data in gray)" 10.What does the green trace at the back of the yellow trace in Fig. 1F represent?As explained in comments #9, the gray trace (instead of green trace) at the back of the yellow line represents the raw fluorescence data, directly extracted from the DNA polymerase trajectory.In contrast, the yellow line illustrates the filtered signal using a Savitzky-Golay filter, as detailed in our methods section.This approach was employed to enhance the clarity and interpretability of the fluorescence signal, facilitating a more precise correlation with the DNAP's mechanical activities.We revised the caption for clarity.
11.The authors should mention the total length of the DNA handle in the main text.
The total length of the DNA handle used in our experiments was 8393 base pairs.We now included this information in the main text of the revised manuscript.
12. Lines 202-204: This sentence should be used somewhere in the beginning to help authors understand the increasing/decreasing basepairs trends in Figure 1 and others.
We agree that providing this explanation earlier in the text will aid readers in understanding the context of our findings.We now incorporate this sentence at an appropriate earlier section in the revised manuscript.

Drs. Avinash Kumar and Yongli Zhang
We thank both the reviewers for their critical and constructive comments.

Reviewer #2 (Remarks to the Author):
This work combines simultaneous confocal fluorescence and force extension techniques to track the activity of bacteriophage T7 DNA polymerase.Both exonuclease and polymerase activity are explored, where the T7 polymerase is labeled with a fluorophore.As in standard force spectroscopy experiments, changes in the fraction of single and double stranded DNA are tracked through the change in the overall length of the template as one is converted into the other.The new technique revealed in this work directly correlates length changes to the location of the T7 as imaged confocally and in kymographs.This enables a careful exploration of the rates of T7 stalling, release and even replacement on the template.While T7 polymerase activity has been widely characterized in force extension experiments, simultaneous imaging and force facilities answering questions that cannot be answered by force spectroscopy alone, including how long each polymerase is bound and active.
Careful study of pauses and bursts of activity reveal that T7 binding and activity is surprisingly stochastic.The same polymerase may rapidly switch between active and inactive states, followed by release.The picture suggested by this data is not of a single, highly processive polymerase, but of a very rapid release and replacement by others from solution.Furthermore, many T7 appear to bind and never initiate polymerase/exonuclease activity.Together this indicates a 'memory' effect, where active polymerase/exonuclease activity is followed by more of the same and switching between the two modes is rarely seen.An intriguing explanation for this is proposed: that in solution, T7 exists in two conformations (and rarely switching), and unbinding T7 leaves the DNA junction in a specific state favorable only to T7 of the same conformation.This is very interesting work that represents a significant advance over previous polymerase experiments and may suggest similar properties for polymerases from other systems.Furthermore, the data appears to be of high quality (though only a single fluorophore is tracked, and labeling is only 60% efficient), and is carefully analyzed.Finally, several controls for photobleaching and diffusion along single and double stranded DNA are shown.With minor clarifications, this paper should be suitable for publication.
Questions: 1. Figure 1 and Methods: Were the tagged T7 filtered after attachment?
We appreciate the reviewer' query regarding the filtration of tagged T7 DNA polymerase.Following the attachment of the fluorophore to the T7 DNA polymerase, the labelled polymerase was filtered.This step ensures the removal of any unbound or excess fluorophores, thereby reducing background fluorescence and enhancing the specificity of our imaging.The details of this process are revised in the 'Fluorescently labelled DNA Polymerase Preparation and DNA Template Construction' section of the Methods.

Figure 1 and Methods: How large is the tag compared to the polymerase?
The tag used in our study is relatively small compared to that of the T7 polymerase.Specifically, the SNAP-tag, which is ~ 20 KDa in size, is fused to the T7 DNA polymerase (approximate molecular weight of ~96 KDa).We now added this value in the method section for clarity.
3. Figure 1 and Methods: What is the temporal resolution of the experiment?The pixel width is ~1 sec, but the methods and results mention that only events of 3 pixels were considered.So only pol exchanges that take longer that 3-5 sec can be seen?This should be discussed in detail.
There might be a bit of unclarity about the temporal resolution of the instrument.The temporal resolution of our image data was determined by the scanning rate of confocal microscopy which results in a pixel width that is approximately 0.3 sec.In our analysis, only the events lasting >3 pixels (~1 sec) were considered for robust analysis, to minimize artifact from diffusive DNA polymerase in solution and ensure the capture of significant binding events.The temporal resolution, therefore, represents a deliberate choice to ensure robust and meaningful data analys and is around 1 sec instead of the assumed 3-5 sec by the reviewer.We rewrote the text to enhance clarity about this point.4. Figure 1: The red box is not explained in the caption -is it for background subtraction?
We apologize for the confusion.The red box in Figure 1 is indeed used for background fluorescence subtraction.We now clarified this in the figure caption.

Figure 2:
There are a high percentage of non-fluorescent events that show exonuclease activity.Could any of this be due to DNA peeling at these high forces over these long times?This is a great point, and we appreciate the reviewer's query in this regard.However, several aspects of our experimental design and observations suggest that the observed events are due to genuine DNA polymerase activities rather than DNA peeling.Firstly, our experiments were conducted under controlled tension conditions, specifically optimized to observe the activities of T7 DNA polymerase.The forces applied (45-50 pN for exonuclease activity and 10-20 pN for polymerase activity) were within the range to promote distinct polymerase and exonuclease activities without inducing DNA peeling.DNA peeling, while a potential occurrence under high tension (> 65 pN) with low ironic strength conditions 4,5 , typically results in abrupt and significant changes in the DNA length, which were not observed in our data.In our experimental design, the method of DNA tethering and buffer conditions with 100 mM NaCl and 3 mM MgCl2, minimizes the likelihood of DNA peeling.
Further, our analysis of the binding lifetimes of DNA polymerase (Figure 2E) indicates that the observed exonuclease activities are consistent with the behavior of T7 DNA polymerase.The binding lifetimes are in line with what is expected for DNA polymerase functioning rather than DNA peeling.
Last, the non-fluorescent events showing exonuclease activity were consistently correlated with the behavior observed in fluorescent events.This correlation strengthens the argument that these events are representative of bona fide DNA polymerase activities.
While the possibility of DNA peeling is a valid concern, the evidence from our controlled experimental setup and the correlation of non-fluorescent events with fluorescent ones strongly supports that the observed exonuclease activities are due to DNA polymerase action.We have now added the following sentences to the revised manuscript to explain this better.
"Notably, the applied forces (45-50 pN for exonuclease and 10-20 pN for polymerase activity), combined with our DNA tethering method and buffer conditions (100 mM NaCl and 3 mM MgCl2), strongly supports that the observed exonuclease activities are due to DNA polymerase action, instead of DNA peeling, which typically occurs under high tension (>65 pN) with low ionic strength conditions. 4,5" 6. Figure 2: In panel E, the stated lifetimes in the caption do not match the values in the figure or in the text.In the caption, the values are distinct, but in the figure, the values are not distinct considering uncertainty.
We thank the reviewer for the careful examination.In the caption, we stated, "The average lifetime for DNA polymerase under exonuclease events is measured to be t = 1.34 ± 0.06 s, and under polymerase events, it is t = 1.11 ± 0.08 s."However, in the figure, we attempted for a representation with one decimal place for clarity and conciseness, thus rounding off these values to 1.3 s and 1.1 s, respectively.The earlier choice was to maintain the figure's readability and avoid overloading it with excessive numerical precision that might not significantly impact the interpretation of the data.We have now reviewed the representations across the figure, caption, and main text to ensure consistency and clarity.The revised values accurately reflect the data and are consistent across all elements of the manuscript.
7. For all figures, please check the panel ordering.It is confusing to follow panels that go in order right to left, then down, then back up to the top row.
We acknowledge that the current ordering of panels might be confusing, and we appreciate your suggestion to improve readability.In the revised manuscript, we revised the panel ordering of these figures.
8. The first equation is equation 3, but shouldn't this be started at 1?
We appreciate your attention to the equation numbering in our manuscript.The numbering of equations starts from the Methods section, where two equations precede the one referenced as Equation (3) in the main text.These two earlier equations ( 1) and ( 2) are critical for establishing the foundational calculations that lead to Equation (3) and thus were referred earlier in the main text.This numbering was chosen to maintain a logical and sequential flow of the mathematical derivations presented.
The authors present single molecule kinetic analysis of DNA polymerization by monitoring changes in length under tension on the template strand resulting from differences between single strand and double stranded DNA.By simultaneously monitoring fluorescence of labeled enzyme and polymerization, the authors describe exchange between polymerases during polymerization.The methods used by the authors are clever and the data appear to be interpreted rigorously.
(1) However, their method is unable to resolve single nucleotide addition steps and is limited to 10 ms per time point.It is not the "high resolution" the authors claim.Moreover, the force on the DNA alters the kinetics as described below.
We agree that a relative term such as "high resolution" is not useful and might even distract from the actual strength of the method.Hence, as suggested, we removed the term "high resolution".
Regarding your concern about the applied force on the DNA template affecting kinetics, we address this point in detail in our response to Comment #4.
(2) There is no description of the formation of a replication fork.The T7 DNA polymerase replication complex has been well characterized defining the coupled action of the helicase, polymerase, and exonuclease with leading and lagging strand synthesis (1).Rather it appears that the authors are only examining DNA elongation with a template and a primer with no duplex DNA and helicase ahead of the polymerase.The current study ignores these detailed studies and presents data that conflict with well-established properties of the polymerase.
We thank the reviewer for this comment.In our approach, we deliberately employed a simplified system to isolate and closely examine the intrinsic exchange dynamics of DNA polymerase independent of helicase action and other replication machinery.This methodological choice was made to specifically explore whether DNA polymerase can exchange autonomously, which is challenging to study in more complex replication systems.Our study does not contradict the established roles and interactions of polymerase, helicase, and exonuclease but rather complements these findings by offering insights into the polymerase's behavior in a controlled environment.
We recognize that this simplification might raise questions about the relevance of our findings to the more complex in vivo replication machinery.To address this, we revised the manuscript to include a clearer explanation of our experimental approach and its rationale in Introduction and Methods.In the Discussion, we also discussed how our findings on the autonomous exchange dynamics of DNA polymerase relate to the established knowledge of T7 replication fork dynamics in the context of error correction by DNA polymerase.We hope this will provide readers with a comprehensive view of the polymerase's behavior both as an independent entity and as part of the larger replication complex.
(3) The most severe limitation of the paper is the lack of quality controls for the changes induced by modification of the DNA polymerase.Most importantly, the authors use T7 gp5 polymerase fused with thioredoxin accessory protein.The only reference given in the text to the use of the fusion protein is a 1987 paper from Richardson's lab, which did not use a fusion protein.Studies have shown that without thioredoxin the DNA polymerase is no longer processive and "undergoes frequent dissociation from and rebinding to the DNA" (2), similar to the result reported in the current manuscript.No controls are given in this manuscript to assess the effect of fusing the thioredoxin to the polymerase.This is a fatal flaw since thioredoxin binding greatly affects polymerase activity and DNA dissociation rates.Fusing thioredoxin to achieve optimal interactions with the flexible DNA binding domain of the polymerase is not a trivial task and would require significant refinement and testing by established methods to define polymerization rates and processivity.A likely explanation of the results presented in this paper is that a less than optimal covalent linkage of thioredoxin to the polymerase was insufficient to stabilize an otherwise flexible DNA-binding domain leading thereby failing to achieve the desired tight DNA binding.
The expression of the T7 pol-thioredoxin fusion protein with an additional SNAP tag (182 amino acids, 19.4 kDa) is also not well described.Was the SNAP tag added to the N-or Cterminus of T7 gp5 and was a flexible linker included?The SNAP tag is likely to introduce a significant change to the polymerase structure and dynamics for which there are no controls.The addition of a large fluorophore further compounds problems.T7 DNA polymerase is a dynamic enzyme where conformational changes dictate nucleotide specificity and proofreading.T7 DNA polymerase has been extensively characterized by single turnover kinetic methods with single base-pair resolution, far exceeding the capabilities of single molecule methods to define reaction kinetics and mechanism (3-9).The results of numerous studies are at odds with the authors' conclusions.The enzyme catalyzes replication at ~300 bp/s and dissociates from DNA at with a rate constant of 0.2/s giving a processivity of 1500 bp.The exonuclease reaction on ssDNA occurs at 1000/s but with duplex DNA the exonuclease is governed by the rate of transfer of the primer strand from the polymerase to the exonuclease site.The kinetic partitioning between polymerization and exonuclease reaction has also been well defined.These studies should stand as a benchmark for evaluating any labeled enzyme to quantify the effects of modifications due to fusing with other proteins and adding fluorescent labels.
For each of these issues, the authors must address the effects of their significant enzyme modifications.Otherwise, their results only apply to the modified enzyme and are not applicable to polymerase function in the cell.At the very minimum, the following experimental measurements are necessary.Single turnover kinetic studies in solution can be easily performed with native and modified enzymes to quantify the effects of modifications.We appreciate the reviewer's insightful comments on the potential impact of the modifications made to the T7 DNA polymerase in our research.Our study employs a modified T7 DNA polymerase, which includes a SNAP tag for fluorescence labeling and a fusion to thioredoxin.These modifications are crucial for single-molecule analysis and were meticulously carried out to preserve the enzyme's functionality.Comparative analyses with other studies, both single-molecule and ensemble approaches, corroborate the validity and significance of our results.See the details about this below.
The fluorescence labelling of T7 DNA polymerase was necessary for our single-molecule analysis.Direct labeling of the polymerase at the C-terminal, N-terminal, or cysteine residues often resulted in inactivity in our trials.Consequently, we opted for an N-terminal SNAP-tag.Although this tag slightly increases the polymerase's molecular weight (by ~20%), the use of SNAP-tag fusion proteins for fluorescence labeling is a widely recognized technique in bioimaging both in vivo and in vitro studies 6,7 .The choice of the labeling site and fluorophore size were carefully considered to minimize interference with the enzyme's active site and conformational changes (see Methods).We also compare the properties of our modified DNA polymerase with that of the commercial polymerase (Figure R1 and Table R1) showing similar results.
The fusion of trx to DNA polymerase aims to preserve DNA polymerase activity after SNAPtag fusion and to avoid any potential inactivity of polymerase.In our study, we cloned gp5 from T7 and trxA gene from E. coli into the pET-Duet1 multiple cloning sites.A flexible linker was included to minimize steric hindrance and maintain enzyme functionality.Our modifications to established protocols 8,9 were designed to maintain the native characteristics of the polymerase while meeting the specific requirements of our experimental setup.The reference to the 1987 Richardson lab paper 9 was to acknowledge the origin of this approach.We intended to also cite this publication from 2003 Richardson lab paper 8 , which demonstrated a covalently linked DNA polymerase with trx showing comparable activity between covalently linked and infusion versions of gp5+trx.We revised our Method section to clarify this.
To ensure the recombined DNA polymerase functioned as close to its native state as possible, we added 625 nM dNTPs (mix) and 2-4-fold excess trx, as supported by previous publications from Richardson lab and Johnson Lab 8,10 .Through these steps, we ensure the fused DNA polymerase retain its native processivity.This information, initially in the raw data metadata, is now detailed in the revised manuscript's Methods section.
Lacking access to radio-labelled nucleotides and detection methods for single turnover assays, we employed single-molecule methods for activity test and comparison, yielding results comparable to published data using the same methods in the same lab.Thus our T7 fused DNA polymerase-thioredoxin (Figure R1   Moreover, as suggested by the reviewer, we compared our single-molecule data with ensemble research and other single-molecule studies to ensure consistency and reliability (see Table R1).Our replication rate, while not identical to previous findings, aligns well within expected ranges.Direct observation of on-rate and off-rate provided a unique assessment of processivity.The slight differences in these values may be attributed to variables such as template length, applied tension, and solution conditions (e.g., ion strength and crowding agents).Current study 30nM 100~350 bp/s with tension 0.75-0.9s -1

Current report
As a final note, in this report, our primary objective was to correlate kinetic activity (measured through changes in DNA template length) with real-time fluorescence monitoring.This method, demonstrated in our report, can be beneficial for many other single-molecule studies investigating DNA-protein interactions.The conclusions drawn in fact align well with recent findings on DNA replication in T7 replisomes [13][14][15] , bacteria [16][17][18][19][20][21][22] , and eukaryotes 23,24 .
(4) The effects of force on the DNA may be responsible for some of the unusual behavior seen in this report.Structural studies show that the template DNA enters the active site at 90 degrees to the growing duplex (10).Therefore, as suggested in the present studies and many previous reports using this method, adding tension to the two ends of the DNA distorts the polymerase active site.It is remarkable that the authors show that a force of 40-50 pN disrupts the polymerase activity so that only exonuclease reaction is seen.But they assume that dropping the force only 2-4-fold to 10-20 pN allows measurement of polymerization without affecting fundamental kinetic parameters including the polymerization and DNA dissociation rate.The authors' analysis rests on an unsupported assumption, and it is likely that the unique conclusions put forth by the authors regarding fast dissociation of the polymerase are an artefact of their methods and enzyme preparation, and there are no controls offered to overcome this objection.
We appreciate reviewer's remarks regarding the potential force effects on the DNA substrate and its implications for our observations.Firstly, while our study applies an external force to the DNA substrate, it's important to note that in cellular environments, DNA template is also influenced by tension 25,26 .This tension can be induced by protein-bound events or structural changes within the DNA.For instance, when a mismatch is incorporated into the DNA duplex, it can induce tension/stress at the replication junction 27,28 .Such tension could facilitate the transition of DNA polymerase to function in an exonuclease mode for proofreading 29 .Such forces on DNA template are challenging to study with conventional methods.Our approach, therefore, although simplified, attempts to mirror certain aspects of the dynamic mechanical environment that DNA polymerase encounters in vivo.
Regarding your concern about the predominance of exonuclease activity at higher forces (~50 pN) and polymerization activity at reduced forces (10-20 pN).This effect is already known and reported for a long time.For instance, Wuite et al. demonstrated similar behavior in T7 DNA polymerase under mechanical forces 1 .Worth mentioning, these force-dependent activity is not specific to T7 DNA polymerase, studies on T4 DNA polymerase 30 , E. coli DNA polymerase 31 , mitochondrial DNA polymerase 32      29 show similar results.Our measurements of DNA polymerase binding duration at higher (i.e.:50 pN) and lower (i.e.: 20 pN) tension revealed comparable durations and dissociation rates with earlier studies (Figure 2E).Moreover, it is essential to note that while the applied force is an external factor, it does not necessarily distort the enzyme's active site directly.The force primarily affects the DNA template's conformation, which in turn could influence enzyme activity indirectly.The ranges of forces we used were chosen based on established literature 1,12,[29][30][31][32] .
We acknowledge that the kinetics observed in our study, under the influence of applied force, may differ from the enzyme's behavior in a single turnover assay and we recognized that both methods are not the same as the natural cellular environment.However, our observations provide valuable insights into how tension, whether externally applied or intrinsically generated, can influence DNA polymerase dynamics.This aspect of our study adds to the understanding of DNA polymerase functionality under varying mechanical conditions, which is pertinent to both in vitro and in vivo scenarios.Plus, the potential differences in kinetics observed in our study compared to ensemble biochemistry studies without tension offer valuable information about the enzyme's adaptability and mechanosensitivity.
We presented these points clearer in our revised manuscript to ensure that the implications of our findings are appropriately contextualized within the broader framework of DNA replication dynamics, particularly emphasizing the role of tension in regulating DNA polymerase activity.
(5) Memory effects have been proposed from single molecule kinetic studies, but to my knowledge there has never been definitive data to support such postulates.Enzymes do not have memories; they are governed by rate constants for transitioning between states.
We recognize the reviewers' concerns about the "memory effect.Indeed, enzymes, being non-sentient entities, do not have memories in the conventional sense.However, what we describe as a 'memory effect' is a metaphorical way of interpreting the observed phenomena where the behavior of DNA polymerase upon rebinding at the replication fork appears to maintain its previous state (either engaged in polymerization, exonuclease activity, or a paused state).Our data revealed that the catalytic activity of DNA polymerase is preserved over a longer time than the actual binding time of a single protein molecule, leading to the chemically continuous yet kinetically discontinuous nature of replication.We now further clarify this in the revised manuscript.

Response to Reviewers' Comments
Reviewer #1 (Remarks to the Author): The authors have made careful revisions to the manuscript and addressed our concerns.We recommend its publication in Nature Communications.
We thank the reviewers again for their effort and careful evaluation of our manuscript.
Reviewer #2 (Remarks to the Author): The thoughtful response to our questions includes many key details that will significantly enhance this manuscript for both the general reader and the specialist.The results represent a significant advance in our understanding of polymerase dynamics.
We thank the reviewers again for their input and constructive suggestions to improve our manuscript.

Reviewer #3 (Remarks to the Author):
This manuscript does not present any new information and does not meet the standards for rigorous biochemical analysis.Details are given in the attached pdf.In their response to the prior critique, the authors have present little new data, only arguments that do not adequately address the fundamental flaws of the paper.
To address the concerns of the reviewer we have taken great care to revise our manuscript and now include a separate Extended Materials document, describing the construction of the pKYB1 DNA and the preparation of the tagged SNAP-DNA polymerase, to provide the detailed information requested.In the revised manuscript, we also explicitly acknowledge the limitations of our resolution in detecting single-nucleotide events.We have made it clear that our study utilizes recombined SNAP-DNA polymerase, detailing the rationale behind this choice and its relevance to our research objectives.To address concerns regarding the structural integrity and functionality of our recombined DNA polymerase, we now include a comparison between the predicted structure of SNAP-DNA polymerase and experimentally determined T7 DNA polymerase structures.To validate the functionality of the SNAP-DNA polymerase, we conducted comparative analysis using two independent assays: real-time DNA primer extension assay and single-molecule assay.Our point-topoint responses below are in blue and changes in the text since last round of revision are in red.
1.The authors inaccurately present this work as characterization of polymerization at a replication fork.They have not described the template/primer system they used, and all evidence points to the use of a single template and primer in the reaction, not a replication fork which is much more difficult to assemble and study.To present this as studies at a replication fork is misleading and dishonest.
Indeed, in our study, we employed a DNA construct that mimics parts of the replication fork.The DNA construct is prepared with biotin labels at both ends, based on established protocols 1 .See Figure R1A (also in Figure 1A of the manuscript, also in Extended Materials for detailed procedures) of the schematics of our DNA construct.Our construct includes a 25 nt 5'-end overhang to enable exonucleolysis, thereby creating a ss/dsDNA junction for polymerization under controlled tension conditions (Figure R1B, also in Extended Figure 1).
We agree with the referee that 'replication fork' doesn't describe our assay properly and we now corrected the terminology "DNA replication fork" throughout the manuscript, and made it clear in our Method section that our DNA construct mimicks part of a replication system.
Figure R1.Schematic of the DNA template (Figure R1A) and sequence of PKYB 1 construct (Figure R1B) used in the current study.The design of this DNA construct can be referred to publications 1 and Method section in the manuscript.The green letter in panel B indicated biotinylated dATP used for bead tethering.
2. This paper has nothing new to offer.In response to the criticisms of their methods, the authors present a table comparing the estimates of the rates of polymerase dissociation from the DNA to rates published in 1993.Although this comparison may be used to justify the validity of their methods, their errors are much larger than estimates published in 1993.Most importantly, this comparison demonstrates that there is nothing novel in the current publication.
We respectfully disagree with the reviewer's assessment in this regard.The referenced polymerase dissociation rate of 0.2 s -1 from studies published in 1993 was actually suggested by the reviewer in previous comments as a benchmark.Our aim in comparing the dissociation rates obtained in our study (0.75-0.9 s -1 ) with those from both ensemble research (0.2 s -1 ) as well as other single-molecule studies (0.6-1.3 s -1 ) 2,3 was to validate the accuracy and reliability of our measurements and analytical methods.This comparison is crucial for demonstrating that our single-molecule approach, despite of using fluorescently labelled protein, aligns well with established data.
Furthermore, the measured dissocation rate (0.75-0.9 s -1 ) across various force ranges (20-50pN) demonstrate a clear understanding of the dissociation dynamics of DNA polymerase under different tensions, contrary to the assertion that this range of rates represent a large error as suggested by the reviewer.We acknowledge the reviewer's concern regarding the novelty of our publication.However, we emphasize that our study's novelty lies in the application of a correlative single-molecule methodology to explore the dynamics of polymerase exchange in a more detailed and nuanced manner than previously possible.
3. The authors justify the use of their method based on the scores of publications using these techniques in the past two decades.The methods were limited when first presented and those limitations have not been rectified in the intervening years.The methods have limited time resolution and are limited by the inability to measure single nucleotide incorporation events.The authors have not clearly stated the limits of resolution in their measurements of DNA polymerization.Similarly, artifacts due to the addition of the SNAP tag cannot be brushed aside just because everyone else does its without appropriate controls.
We acknowledge the limitations mentioned, specifically the temporal resolution constraints and the challenge in detecting single-nucleotide incorporation events directly.Our experimental setup, combining optical tweezers with fluorescence microscopy, offers a temporal resolution (~0.3s/scan) that, while not sufficient to resolve individual nucleotide incorporations, is highly effective in capturing the rapid exchange dynamics of DNA polymerase and its overall activity patterns.To address the reviewer's comments, we have added the following text to the manuscript to clarify the resolution limits of our experimental setup: "The temporal resolution of our correlative optical tweezers-fluorescence microscopy setup is determined by the confocal scanning rate and the signal acquisition time required for reliable fluorescence detection.While this resolution does not allow for direct observation of single nucleotide incorporation events, it is optimized to capture the rapid exchange dynamics of DNA polymerase and polymerase activity patterns during replication, complementing higherresolution techniques." To address the concerns about the effect of SNAP tag fusion and subsequent fluorescence labelling, we first investigate the structural integrity of our recombined DNA polymerase by employing the RoseTTAFold Modeling Method 4 to predict the structure of our utilized SNAP-DNA polymerase.Aligning this model with the experimentally determined structure of T7 DNA polymerase (PDB:1T7P) and a DNA primer/template complex allowed us to verify that the structural integrity and active site functionality is expected not to be impacted.This analysis, presented in Figure R2 and Extended Figure 3, shows that the SNAP-tag is out of the way and should minimally impact the polymerase's functionality.(A) Predicted structure of SNAP-DNA polymerase using the RoseTTAFold method 5 , aligned with the experimentally determined T7 DNA polymerase (PDB:1T7P) 4 to model the complex with DNA primer/template.Color: SNAp-tag in magenta, N-terminal with flexible GS-linker in red, and gp5 protein in cyan.(B) Comparison of the RoseTTAFold-predicted SNAP-DNA polymerase structure with T7 DNA polymerase (PDB:1T7P), demonstrating preserved structure and active site for DNA binding.The experimental T7 polymerase is in green, with trxA indicated in light green.(C) Close-up of the N-terminal with a GS-linker and SNAP-tag, illustrating the tag's distance from the active site, suggesting minimal impact on binding and activity.(D) Zoomed-in view of the trxA-binding domain, showing its accessibility for trxA interaction.
We also conducted control experiments, using real-time DNA primer extension assay 6 and single-molecule assays 3 , showing that our SNAP-tagged polymerase has comparable activity to commercial T7 DNA polymerase (NEB, # M0274L) (Extended Materials, Figure R3, also in Extended Figure 4).In particular, using a real-time DNA primer extension assay, we compared DNA polymerase activity across a range of enzyme concentrations.The assay               fluorophores at various DNAp concentrations.Our results demonstrate that both the commercial T7 DNA polymerase (Figure R3A, shown with a gradient of blue) and our SNAPtagged polymerase (Figure R3A, shown with a gradient of green) displayed similar activities under our measured concentrations.Further, at the single-molecule level, we compared the activity of both polymerases under various tensions.This comparison showed that the SNAP-tagged polymerase's (Figure R3B) kinetics closely match those of the published data (see Figure R3A) 3 .These findings indicate that the SNAP tag does not significantly impact the enzyme's activity or DNA interaction.As suggested, we have also refined the manuscript to explicitly state that our findings are based on a recombinant DNA polymerase with a SNAP-tag.The manuscript now includes the following statement: "In this study, we used a recombinant DNA polymerase with a SNAP-tag for protein labelling.Future research should consider direct labelling methods for the polymerase, although attempts to do so were not successful in our case." 4. In response to the prior criticism regarding use of a fusion between the polymerase and the thioredoxin cofactor, the authors state now refer to a 2003 paper by Johnson and Richardson and they blur the line between a fusion and a covalent disulfide crosslink.While Richardson paper demonstrates that the crosslink positions the thioredoxin appropriately for activity, this does not address at all whether fusing the thioredoxin to the N-or C-terminus is effective.
We appreciate the reviewers' insightful comments and the opportunity to clarify our experimental approach.Upon re-examination of our protein sequence, laboratory documents, and experimental data, we recognized an oversight in our initial design.Our prior description mistakenly suggested the successful expression of a thioredoxin fused directly to the T7 DNA polymerase.However, detailed examination of our protein sequence and laboratory documentation revealed that attempts to create a trxA-gp5 fusion protein were unsuccessful.This was due to the presence of stop codons immediately following the gp5 gene, which led to premature translation termination.This was conclusively identified through the use of the Expasy Translation Tool for predicting protein translation (Figure R4A) and confirmed by SDS-PAGE.The observed molecular weight was consistent with the absence of a trxA-tagged component, as shown in Figure R4B and detailed in the Extended Materials.
The figure at the right shows the structure of T7 DNA polymerase (gray) with thioredoxin in cyan.The N-and C-termini of the polymerase are shown in blue and red spheres, respectively.Attaching thioredoxin to either the N-or the C-terminus is not tenable, even with a very long linker.
[redacted] Our initial intent was inspired by the potential to enhance T7 DNA polymerase's processivity through thioredoxin fusion.Addressing the reviewer's concern, we acknowledge the practical challenges of effectively attaching thioredoxin to either the N-or C-terminus of the polymerase, even with the use of a lengthy linker.In our study, the addition of an excess amount of free thioredoxin-a common practice for working with T7 DNA polymerase 1,13 resulted in normal enzymatic activity even though the fusion was not successful.Thus, this methodological clarification does not alter the interpretation of our data, or the conclusions drawn from our study and in fact might be a blessing in disguise.We have amended the Methods section of our manuscript to accurately reflect these findings and clarify the experimental design.We regret any confusion our initial submission may have caused and are grateful for the chance to correct this error.
5. This and other publications using single molecule methods have overstated the significance of DNA polymerase pausing.No one has shown that the pausing is not an artefact caused by the methods of analysis.In any event, the fraction of polymerase molecules in a paused state is insignificant.Therefore, the one novel contribution of these methods, as applied to DNA polymerization, is unimportant and not original with this paper.
We respectfully disagree with the assertion that the significance of DNA polymerase pausing is merely an artifact of analysis methods or is insignificant.Firstly, we employed two independent methods (mechanical force and fluorescence microscopy) to analyze and interpret our data, ensuring that the observed pausing events are not artifacts of a particular analysis technique.See example pausing events detected in our study, from mechanical measurement (Figure R5A, red arrow) and fluorescence measurements (Figure R5B, red arrow).This multi-faceted approach strengthens the validity of our findings, demonstrating that pausing is an inherent feature of DNA polymerase dynamics rather than a methodological artifact.Contrary to the reviewer's assertion, our data indicate that pauses are not merely stochastic interruptions but are integral to the polymerase's function, possibly allowing for exchange of DNA polymerase and processivity regulation (Figure R5C and R5D).This aligns with emerging evidence suggesting that transient pausing can influence enzymatic activity and genome stability.In the current study, pausing events are captured via two independent methods: optical tweezer mechanical measurements (Figure R5A) and confocal microscopy imaging (Figure R5B).Correlation of these datasets identifies replication segments with pauses, highlighting a prevalence of single-type burst segments (Figure R5C) and transitionaltype burst events (Figure R5D).
Furthermore, it is crucial to recognize that DNA polymerase pausing has been documented across various studies with various methods, attributed to a multitude of factors such as DNA secondary structures 7 , the presence of DNA lesions 8,9 , specific sequences 10 and replication-transcription interaction 11 .In our study, we extend the understanding of DNA polymerase dynamics by suggesting that dissociation events could contribute to observed pausing.Contrary to the assertion that the pausing is "insignificant" and our contributions "unimportant and not original," we argue that elucidating the underpinnings of polymerase behavior, including pausing, enriches our comprehension of the replication machinery's adaptability and error-correction mechanisms.
Finally, our work extends beyond merely identifying pausing events.We offer new insights into the context and implications of these pauses, particularly in relation to the enzyme's overall activity and replication fidelity.This is supported by a detailed analysis of pause frequency, duration, and correlation with enzymatic activity changes, providing a more comprehensive picture of DNA polymerase behavior than previously reported.
6.There is little doubt that the tension on the DNA introduces artefacts by perturbing the structure of the enzyme.At the very least the authors need to make measurements as a function of force and extrapolate to zero force.
The impact of force on DNA polymerase activity has been studied before and discussed in quite some detail 2,3 .But as suggested, we conducted additional experiments across a range of applied forces, including analysis at low tentions at 10pN to asses the potential impact of applied tension on the DNA structure and, consequently, on the observed behavior of the DNA polymerase (see Figure R2B, also in Extended Figure 4).That said, for exonuclease (exo) activity, data has shown that this reaction appears largely independent of the applied force.This independence from force underscores a fundamental aspect of DNA polymerase's function-its ability conduct proofreading without being significantly influenced by the mechanical state of the DNA.Such a characteristic is essential for the enzyme's role in maintaining genomic integrity across various physiological conditions.Moreover, it is not very likely that DNA polymerase inside cells/bacteria will process DNA that is under zero tension, given the many contrains on the DNA inside the crowed interior and it being continously processed by many proteins.Thus extropolating to zero force will not provide more insight that the force range that we considered.In this manuscript, the authors have employed correlative fluorescence microscopy and force spectroscopy to elucidate the dynamics of the T7 DNA polymerase gp5 during its exonuclease and polymerase activities.Their study uncovers a rapid polymerase exchange phenomenon in the presence of the processivity factor thioredoxin, independent of the complete replisome assembly.Additionally, they report a 'memory effect' influencing polymerase exchange, which they attribute to the state of the single-stranded/double-stranded DNA junction.
The manuscript has undergone prior review by three referees, with the first two expressing satisfaction upon revision.The third reviewer, however, persisted with concerns.Initially, the critique centered on the potential non-representative nature of the gp5-thioredoxin-SNAP-tag fusion and the possible influence of applied force on DNA affecting the gp5 activity.The authors have since addressed these points satisfactorily in their revision.
Subsequently, the third reviewer raised issues regarding the novelty of the findings, suggesting that polymerase exchange has been observed at authentic replication forks involving helicase-mediated DNA unwinding.I find the insights provided by the current work, particularly the independence of polymerase exchange from helicase interaction and the noted 'memory effect,' to be significant contributions.The third reviewer's insistence on single-nucleotide resolution overlooks the value of single-molecule visualization techniques in revealing enzymatic reaction heterogeneities and kinetics, despite lower spatial resolution.
The thorough critique from the third referee has led to an important realization that the gp5 construct used lacked a thioredoxin fusion.But, it seems that the authors had also included gp5 in their experiments.Overall, I feel that the authors have addressed the comments from the third reviewer, enhancing the quality of the manuscript.I would advise the authors to consider the remaining remarks of the third reviewer, particularly pertaining to Extended Data Figure 4.The concerns about novelty should not overshadow the merits of the study.Once these final points are addressed, I advocate for the manuscript's publication in Nature Communications.

Figure 2 :
Figure 2: In panel E, the stated lifetimes in the caption do not match the values in the figure or in the text.In the caption, the values are distinct, but in the figure, the values are not distinct considering uncertainty.

8 .
In the kymographs (Figs.1E, 3A, 4D and others), what does the high intensity at the bottom and top represent?In Figs.4A and 4B, the authors indicate that the top and bottom intensity bands are attributed to signals from bead? Can authors clarify it and mention it in all the kymographs?
a. What is the active site concentration of the modified, labeled enzyme?That is, what fraction of the enzyme still binds and extends DNA? b.What is the rate of polymerization in solution?c.What is the Kd for DNA binding to the altered enzyme?d.Most importantly, what is the rate constant for DNA dissociation?
C&D) displays kinetics that is more or less the same as the commercially available DNA polymerase of the published data (see Figure R1 A&B) 3 .

Figure R1 .
Figure R1.Comparative Analysis of Enzymatic Activity Between Commercial T7 DNA Polymerase (Control) (panels A and B), adapted from ref3  and Modified DNA Polymerase (Experimental) (panels C and D).(A) and (C): These panels illustrate the polymerization and exonucleolysis activities of the commercial and modified DNA polymerases, respectively.(B) and (D): These panels depict the relative probability of DNA polymerase binding with its exonuclease (exo) active site for both the commercial and modified enzymes.

Figure R2 .
Figure R2.Structure and Alignment of Predicted SNAP-DNA Polymerase with T7 DNA Polymerase.(A)Predicted structure of SNAP-DNA polymerase using the RoseTTAFold method5  , aligned with the

Figure R3 .
Figure R3.SNAP-DNAp exhibits activity comparable to commercial DNAp.(A) Analysis of DNA polymerase activity using a real-time DNA primer extension assay reveals comparable performance between commercial T7 DNA polymerase (illustrated with a gradient of blue) and SNAP-DNAp pre-mixed with trx at a 1:4 ratio (shown with a gradient of green), across a range of enzyme concentrations (2.3nM, 4.7nM and 9.4nM).                 Data were plotted with three independent measurements.(B) Polymerase activity was quantified by analyzing the initial linear phase of the fluorescence intensity decrease (first 3 minutes).Values and errors (sem) were derived from three independent experiments.Furthermore, we compare the DNAp activity using singlemolecule assay (C and D).Comparative Analysis of Polymerization and Exonucleolysis Activity Between

Figure R4 :
Figure R4: Analysis of SNAP-DNA Polymerase Translation Purification.(A) Protein translation prediction for the pETDuet-1_SNAP-DNA polymerase plasmid, using the Expasy Translation Tool from SIB Swiss Institute of Bioinformatics.Results show that trxA-tag fusion is disrupted by stop codons post-gp5 gene, with only the DNA polymerase's open-reading-frame successfully expressed (highlighted in red).(B) SDS-PAGE gel electrophoresis confirmed the purified product as solely SNAP-DNA polymerase, with the trx component absent.

Figure R5 .
Figure R5.Representative Pausing Events and Integrated Replication Process.In the current study, pausing events are captured via two independent methods: optical tweezer mechanical measurements (FigureR5A) and confocal microscopy imaging (FigureR5B).Correlation of these datasets identifies replication segments with pauses, highlighting a prevalence of single-type burst segments (FigureR5C) and transitionaltype burst events (FigureR5D).
7. The authors' opening statements that there is very little known about DNA polymerization does not reflect the state of the field.The mechanistic detail now known about the polymerase kinetics beyond the limits of single molecule methods.It appears there has been a misunderstanding about the phrasing of our manuscript's introduction.Our manuscript states, "Despite extensive studies on DNA replication, the exchange mechanisms of DNA polymerase at the replication fork (now changed to 'during replication') remain unclear."Our intent was not to suggest a general lack of knowledge about DNA polymerization mechanisms, which indeed are well-characterized through extensive biochemical, structural, and mechanistic studies.Instead, our focus was specifically on the exchange mechanisms of DNA polymerase at the replication fork, an area that remains less understood, especially in the context of dynamic, real-time processes observable through single-molecule techniques.This distinction is crucial, as the rapid and autonomous exchange of DNA polymerase at the replication fork, as observed in our study, underscores complex regulatory mechanisms that govern replication fidelity and efficiency in a cellular context.Our findings, as detailed in both the manuscript and the response to the first round of revisions, highlight these dynamic processes, contributing to the broader discourse on DNA replication mechanics.
Figure 1A.Simulation to mimic the authors' data.